We thank Rayhan Arif and Hassaan Shahawy for camera work, Tripti Gupta and Frederike Alwes for assistance refining the cell ablation technique, and Extavour lab members for feedback on the data, video and manuscript. This work was partially supported by the Harvard Stem Cell Institute (Seed Grant number SG-0057-10-00) the Ellison Medical Foundation (New Scholar Award number AG-NS-07010-10) to CGE, and Harvard College Research Program awards to ARN.

Comments that may be helpful in executing certain steps are indicated in italics.
1. Day 1: Preparation of Materials
<ol>
	<li>Prepare the following materials (see Table of Materials and Equipment):
	<ul>
		<li>15 cm and 22 cm Pasteur pipettes</li>
		<li>Diamond scribe</li>
		<li>filtered artificial seawater (salinity between 0.0018-0.0020) containing 1 mg/ml Amphotericin B (1:100 of a 100 mg/ml stock solution), 100 units/ml penicillin and 100 mg/ml streptomycin (1:50 of a stock solution containing 5,000 units/ml of penicillin and 5,000 mg/ml of streptomycin)</li>
		<li>filtered artificial sea water (salinity between 0.0018-0.0020) without additives</li>
		<li>a large plastic container for housing couples (at least 15 cm x 15 cm x 5 cm)</li>
		<li>small (3 cm or 5 cm) Petri dishes lined with Sylgard</li>
		<li>small (3 cm or 5 cm) Petri dishes</li>
		<li>a watchglass or glass multi-well plate</li>
		<li>forceps (we use blunt forceps derived from old Dumont #5 forceps that were accidentally damaged in other laboratory procedures, which we have repurposed by using a forceps sharpening stone to ensure that the ends of the forceps are smooth, that both tines are the same length, and that they no longer have sharp points. Extremely fine forceps, such as new Dumont #5 forceps, are undesirable as they may injure embryos and/or females.)</li>
		<li>a Pasteur pipette with the end widened (see step 1.2 below)</li>
		<li>an embryo retrieval tool consisting of a thin, curved tungsten wire embedded into the thin end of a Pasteur pipette (could be replaced by an alternative implement; see 1.3 below)</li>
		<li>a mouth pipette with a small opening (see step 1.4 below)</li>
		<li>a glass needle made from a pulled glass capillary (see step 1.5 below)</li>
	</ul>
	No materials toxic to humans are used in this protocol. However, researchers should ensure that they wear gloves or other protective attire as required by the risk management guidelines of their institution.</li>
	<li>To make the widened Pasteur pipette, first score around a 15 cm Pasteur pipette at the point where the pipette begins to narrow with a diamond scribe, wrap the scored region of the pipette in a Kimwipe or paper towel to protect the fingers, and then carefully break off the tip at the score line. Hold the broken end of the pipette over a Bunsen burner flame for several seconds to smooth the edges. The opening should be slightly narrower than the maximum width of the pipette.</li>
	<li>To make the tungsten wire dissecting tool, insert a tungsten wire into the end of a 15 cm glass Pasteur pipette, and hold briefly over a Bunsen burner flame to melt the glass, fixing the wire in place. Use forceps to gently curve the end of the wire into a hooked shape. See Figure 3 for an example of an appropriate shape for this tool.</li>
	<li>To make a small opening for the mouth pipette, pull the thin end of a 22 cm Pasteur pipette into two parts over a flame, and break off the tip of the small end. Insert into the end of the mouth pipette holder.</li>
	<li>Make the glass needle that will be used to puncture the blastomeres of interest during the ablation procedure using a needle puller. A suitably shaped needle is shown in Figure 4. This needle was pulled on a Sutter P97 needle puller using a program with parameters 2x(heat = 566, pull = 100, velocity = 20, time = 250) + 1X(heat = 566, pull = 100, velocity = 100, time = 250). The specific program and instrument used to make the needle can vary, but the needle must have a fine point, yet also be sturdy enough not to break when pushed against the chorion of the embryo.</li>
</ol>
2. Day 1: Gathering Parhyale&#160;Mating Couples
<ol>
	<li>Use the widened Pasteur pipette (step 1.2 above) to collect mating P. hawaiensis couples from the large culture container, and transfer them to a separate container containing clean artificial seawater. It is not necessary that this seawater be filtered. It is best to collect mating pairs in the later afternoon and leave the couples at room temperature (20-23 &#186;C) overnight, so that the next morning, most embryos will have been recently fertilized and deposited, and thus at early cleavage stages suitable for ablation. Gather at least 20 mating pairs to ensure that some females will be carrying early cleavage stage embryos by the following morning.</li>
</ol>
3. Day 2: Gathering Parhyale&#160;Embryos
<ul>
	<li>The next morning, infuse filtered artificial seawater (FASW) with CO2. Some of the females will have swum free from their males during the night; collect these separated females and anesthetize them in FASW / CO2. The remaining unpaired males can be removed from the mating pair container and returned to the large culture. Remaining mating pairs can be saved for embryo collection later in the day or the next day.</li>
	<li>Transfer a single anaesthetized female carrying embryos to a watch glass in FASW / CO2. All separated females will likely have deposited eggs, which will be visible by eye as pale pink or purple spheroids through the transparent coxal plates of the females (Figure 1A). To determine if the embryos are at early cleavage stages and thus appropriate for blastomere ablation, the researcher will have to remove the embryos from the brood pouch and examine them under a microscope.</li>
	<li>Viewing the procedure under a dissecting microscope, grasp the coxal plates along one side of the body with forceps. Embryos will be visible in the ventral brood pouch between these coxal plates (Figure 1B).</li>
	<li>Insert the thin, curved wire tool (Figure 3; step 1.3 above) into the posterior end of the brood pouch, and gently lift the wire towards the anterior end of the brood pouch to separate the brood pouch coverings (these are modified ventral appendages called oostegites30), opening the pouch and loosening the embryos. Embryos that are within the first hour or so of being laid are all enclosed within a very thin, transparent membrane that will easily be disrupted by the wire tool; this membrane disappears by approximately 2-3 hr after egg laying, and embryos are not bound to each other or to the female in any way from this point onwards throughout embryogenesis. If researchers find it inconvenient or difficult to maneuver the curved wire tool to remove embryos from the brood pouch, an alternative implement may be used, as long as the movements are gentle and no harsh pressure is applied to the embryos or the brood pouch. Other tools appropriate for removing embryos from the brood pouch include blunted forceps or a glass capillary with a sealed and smoothed end.</li>
	<li>Use an unaltered 15 cm Pasteur pipette to flush water through the opened brood pouch, pushing out the embryos, which should settle to the bottom of the watch glass. Transfer the female to clean FASW (without CO2) to allow recovery before reintroducing her to the main culture. Multiple females can be placed into the same recovery dish, but allow females to recover fully before returning them to the main culture, as they may be eaten by other animals if they are still partially anesthetized.</li>
	<li>Use an unaltered 15 cm Pasteur pipette to transfer the embryos to a Petri dish with clean FASW, and view under the microscope to determine their embryonic stage. Separate embryos that are at the third cleavage (stage 4&#160;5), which are suitable for this procedure. It should also be possible to apply this protocol to any cleavage stage prior to germ disc formation (stages 7-8&#160;5).</li>
</ul>
4. Day 2: Ablating Single Cells
<ol>
	<li>Fill Sylgard-lined Petri dish with a shallow layer of FASW. Use an unaltered 15 cm Pasteur pipette to transfer a single embryo to the plate.</li>
	<li>Using blunt forceps, gently roll the embryo to identify the cell to be ablated (Figures 2A and&#160;2B). The germ line precursor g can be identified as the smallest micromere, which sits on top of the smallest macromere Mav. Additionally, g does not directly contact the cell en, which is the micromere directly across from it, as the mesoderm precursor micromeres ml and mr share a cell border in the middle of the embryo. mr is the blastomere to the right of g, and ml is the micromere to the left of g. The sister macromeres of mr, ml, and en are the ectoderm precursors Er, El, and Ep respectively.</li>
	<li>With forceps in the left hand if the researcher is right-handed (or in the right hand if the researcher is left-handed), orient the embryo with the cell of interest to the right (or to the left if the researcher is left-handed), and gently grasp the embryo between the forceps to stabilize it.</li>
	<li>Using a pulled glass needle (step 1.5 above; Figure 4), gently puncture the cell of interest. Do not insert the needle too far, so as not to push the needle into the cells neighboring or below the cell of interest. The needle only has to puncture the chorion and underlying cell membrane, and does not need to extend deeply into the cell to be ablated.</li>
	<li>Exchange the needle used to puncture the embryo for the glass end of a mouth pipette with a fine pulled Pasteur pipette tip (step 1.4 above). Hold the tip of the pipette close to the hole created in the cell of interest, and apply very gentle suction.</li>
	<li>Very gently squeeze the embryo with the forceps. As the content of the blastomere is pushed out of the chorion through the hole created in step 4.4, suck it into the mouth pipette to allow an unobstructed view of the embryo. If the hole is large enough, the nucleus of the punctured cell may be seen to emerge as well. This is a good check to make sure the entire cell contents have been removed and cannot be regenerated.</li>
	<li>Carefully observe the embryo as the blastomere of interest is removed. The border of the punctured cell will recede towards the hole in the chorion, making the intersections of the underlying cells visible. Continue applying pressure until all of the blastomere of interest has been removed. Use forceps to roll the embryo from side to side and visually confirm that all of the blastomere of interest is gone.</li>
	<li>Using an unaltered 15 cm Pasteur pipette, transfer the embryo to clean FASW+ (FASW + 1 mg/ml Amphotericin B, 100 units/ml penicillin and 100 mg/ml streptomycin).</li>
	<li>Raise blastomere-ablated embryos in individual wells of a clean 48-well plate in FASW+, changing 50% of antibiotic-supplemented FASW+ daily. In our hands embryos survive and develop well, even following ablation, at a range of temperatures from 18-28 &#186;C. Researchers should raise their embryos at a temperature for which they have a clean (but not necessarily sterile) surface on which to raise the embryos with minimal disturbance. To be able to compare the timing of developmental events in blastomere-ablated embryos with controls, we refer the reader to the detailed developmental staging information that is available for embryos raised at 18 &#186;C4, 25 &#186;C4, and 26 &#186;C5,6.</li>
</ol>

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The authors have nothing to disclose.

We describe a protocol for manual ablation and complete physical removal of single blastomeres from early cleavage stages of the amphipod P. hawaiensis. We demonstrate use of this protocol by removing the single germ line precursor cell g from an eight-cell stage embryo, and show that ablation has been successful by confirming absence of g&#8217;s daughter cells in later embryogenesis. This protocol can be used to remove any of the micromeres from the embryo at early cleavage st...

The amphipod crustacean Parhyale hawaiensis has emerged over the last decade as a promising model organism with great potential for use in evolutionary developmental biology research1. Among the arthropods, most model systems are insects, and the most extensively studied of these is the fruit fly Drosophila melanogaster. D. melanogaster is a member of the insect order Diptera, and as such displays many embryological features that are derived with respect to those of basally branching insects2. Moreover, insects are nested within the subphylum Pancrustacea3, meaning that insects have their closest relatives within the long-standing &#8220;natural&#8221; group called pancrustaceans, and that this group is paraphyletic. This suggests that in addition to basally branching insect models, studies of other crustaceans are required to gain a broader view of the evolutionary history of the developmental traits and molecular mechanisms that have been so well studied in D. melanogaster. However, very few crustaceans have been well established for experimental laboratory analysis of development. The amphipod P. hawaiensis is a highly tractable laboratory model system, amenable to a range of experimental techniques. Amphipods display many unique features within their parent superorder Peracarida (beach hoppers, scuds, and well shrimps), and are therefore thought to be relatively derived within this group of crustaceans. Nevertheless, the relative ease of embryological and functional genetic manipulation offered by Parhyale make this amphipod a valuable addition to the current inventory of model organisms.
As a laboratory animal, P. hawaiensis offers many advantages. Animals are tolerant to a wide range of temperatures and salinities, and survive well in large cultures of artificial seawater1. It is easy to distinguish between males and females based on clear morphological differences, most notably, the large, hooked, anterior trunk appendages that males use to grasp the females during mating. For embryological and developmental work, P. hawaiensis has several very appealing features. Embryogenesis lasts approximately 10 days and the time to sexual maturity is approximately six weeks at 28 &#186;C (but note that Parhyale survives well at temperatures ranging from approximately 20-30 &#186;C, and that detailed developmental staging information is available for embryos raised at 18 &#186;C4, 25 &#186;C4, and 26 &#186;C5,6). Adults mate all year round in the laboratory, so embryos are available at any time of year. Females lay 2-20 (depending on the age of the female) fertilized eggs into a ventral brood pouch located between the first several pairs of legs (Figures 1A and&#160;1B), and it is possible to gather these embryos very early in development without killing the female or damaging the embryos (Figure 1C). The embryos survive in filtered artificial sea water through to hatching, can be fixed for subsequent gene expression or histological analysis7, and a detailed staging table allows accurate identification of the progress through development5. Robust protocols have been used to perform gene expression analysis by in situ hybridization8-15 or immunostaining4,16,17, functional knockdown by RNA interference13,15 or morpholinos12, and stable germ line transgenesis18. Using the transgenesis system, inducible expression14 and enhancer trap19 methods can also be used to investigate gene function in P. hawaiensis. While a publicly available genome sequence is not currently available, a transcriptome containing transcripts produced during oogenesis and embryogenesis has been de novo assembled and annotated20, and deposited in a searchable database21, facilitating gene discovery. In sum, P. hawaiensis is a highly tractable model organism suitable for multiple experimental and genetic approaches to understanding development.
Unlike the early syncytial cleavages of D. melanogaster, P. hawaiensis embryos cleave holoblastically following fertilization (Figure 2A). Lineage tracing analysis has shown that by third cleavage, each of the third cleavage blastomeres is specifically fated to give rise to one of the three germ layers or the germ line6 (Figure 2B). These data, together with microarray data22, cell lineage analyses6,23, and blastomere isolation experiments4 have suggested that developmental potentials are segregated to at least some third cleavage blastomeres by asymmetric inheritance of cell fate determinants. Accordingly, in blastomere ablation experiments in which the germ line precursor (termed &#8220;g&#8221; in the Parhyale cell lineage nomenclature6) was removed at the eight cell stage, embryos lacked germ cells at later developmental stages4, as indicated by the absence of cells expressing the protein Vasa, which is a germ line marker in most metazoans24. In contrast, somatic blastomere ablation experiments showed that P. hawaiensis embryos also possess significant regulatory capabilities, such that the fates of mesoderm or ectoderm precursor blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres25. How regulative cell fate replacement can occur, and the extent of autonomous cell fate adoption by somatic blastomeres, remains unknown. Experimental embryological techniques such as blastomere ablation can be useful in understanding the relative autonomy and nonautonomy of cell fate decisions26,27&#160;and are therefore of interest in the study of P. hawaiensis embryogenesis.
In the experiments that demonstrated regulative replacement of ectodermal and mesodermal lineages, blastomere ablation was performed by injection28 and subsequent excitation of phototoxic dyes25. While this technique is effective at killing the injected blastomere(s), it does not completely remove the dead cell body from the embryo. In addition, differences have been observed between cell lineage data gathered through to gastrulation stages of embryogenesis by injecting blastomeres with fluorescent lineage tracers28,29, and data gathered by following unperturbed blastomeres through development of the same embryonic stages23. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications.
We previously published the results of cell lineage analyses of embryos in which single cells were manually ablated23. However, the delicate operations required to remove single blastomeres from early cleavage stage embryos have not yet been fully described. Here we present a protocol for collection of P. hawaiensis embryos and manual ablation of a single blastomere from an eight-cell stage embryo. The goal of this method is to achieve complete removal of the cell body from the embryo, allowing observation of the cellular behaviors and cell fate competencies of the remaining cells during embryogenesis and post-embryonic development. Our protocol shows removal of the germ line precursor g (Figure 2C), but can be applied to any cell at the eight-cell stage, or to blastomeres of earlier cleavage stages. In principle, this protocol could be applied to remove single cells from early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

<table><tbody><tr><td>15 cm Pasteur pipette</td><td>Wheaton</td><td>53499-630 VWR</td><td>For transferring intact and blastomere-ablated embryos from dish to dish.</td></tr><tr><td>22 cm Pasteur pipette</td><td>Wheaton</td><td>53499-632 VWR</td><td>For making mouth pipette tips.</td></tr><tr><td>3 cm Petri dishes</td><td>Thermo Scientific</td><td>25382-334 VWR</td><td>Use some unaltered to collect and store embryos; coat some with a layer of Sylgard (see below) to create a working surface for blastomere ablations.</td></tr><tr><td>48-well Plates</td><td>Greiner Bio-One</td><td>82051-004 VWR</td><td>For culturing embryos following ablation.</td></tr><tr><td>Bottle top filters 0.2 &amp;micro;m</td><td>Nalge Nunc International</td><td>28199-296 VWR</td><td>For creating FASW (See below).</td></tr><tr><td>Bunsen burner</td><td>VWR</td><td>89038-530 VWR</td><td>For creating tungsten wire tool and fine tip of mouth pipette.</td></tr><tr><td>Diamond scribe</td><td>Musco Sports Lighting</td><td>52865-005 VWR</td><td>For creating wide-mouthed Pasteur pipettes for collecting couples.</td></tr><tr><td>Forceps</td><td>Fine Science Tools</td><td>11050-10 Fine Science Tools</td><td>For holding anaesthetized females during removal of embryos from brood pouch. These do not have to be fine-tipped Dumont forceps - the tips can be at least as blunt as those of the forceps for which the catalogue number is listed here.</td></tr><tr><td>Fungizone-Amphotericin B</td><td>Sigma</td><td>A2942 Sigma Aldrich</td><td>Add to FASW to a final concentration of 1 mg/ml, and use this solution to culture blastomere-ablated embryos.</td></tr><tr><td>Glass capillaries 4 in long, 1/0.58 OD/ID (mm)</td><td>World Precision Instruments</td><td>1B100-4 World Precision Instruments</td><td>For making needles for puncturing the blastomere to be ablated.</td></tr><tr><td>Glass watchglass 300 mm diameter</td><td>Electron Microscopy Sciences</td><td>70543-30 Electron Microscopy Sciences</td><td>For use in removing embryos from the brood pouch of anesthetized females.</td></tr><tr><td>Instant Ocean Artificial Sea Water (ASW)</td><td>Instant Ocean</td><td>IS160 Aquatic EcoSystems</td><td>Make ASW by dissolving salt in deionized water to a salinity of 1.022-1.024.</td></tr><tr><td>Instant Ocean Filtered Artificial Sea Water (FASW)</td><td>Instant Ocean</td><td>IS160 Aquatic EcoSystems</td><td>Use 0.2 &amp;micro;m filters to sterilize ASW. Make up in small amounts (&amp;lt; 250 ml) as needed.</td></tr><tr><td>Kimwipes</td><td>Kimberly-Clark</td><td>21905-026 VWR</td><td>For safely breaking off the fine end of Pasteur pipettes modified for collecting couples.</td></tr><tr><td>Mouth pipette adaptor</td><td>Drummond Labware</td><td>A5177-5EA Sigma Aldrich</td><td>Insert the fine pulled Pasteur pipette tip into this end, to be used for removing extruded cell contents during the ablation procedure.</td></tr><tr><td>Mouth pipette tubing</td><td>Aldrich</td><td>Z280356 Sigma Aldrich</td><td>Cut this to be long enough to comfortably hang around your neck during the ablation procedure.</td></tr><tr><td>Needle Puller</td><td>Sutter Instrument Company</td><td>P97 Sutter Instrument Company</td><td>For making needles from glass capillaries for puncturing the blastomere to be ablated.</td></tr><tr><td>Penicillin-Streptomycin Solution</td><td>Mediatech</td><td>45000-650 VWR</td><td>Add to FASW to a final concentration of 100 units/ml penicillin and 100 mg/ml streptomycin, and use this solution to culture blastomere-ablated embryos.</td></tr><tr><td>Rubber bulb</td><td>Electron Microscopy Sciences</td><td>100488-418 VWR</td><td>For using Pasteur pipettes to transfer embryos from dish to dish.</td></tr><tr><td>Sylgard 184</td><td>K. R. Anderson, Inc.</td><td>NC9659604 Fisher Scientific</td><td>Make up according to manufacturer&#039;s instructions. Pour a layer 2-5 mm thick into a 3cm petri dish to create a working surface for blastomere ablations.</td></tr><tr><td>Tungsten wire 0.004 inches diameter</td><td>A-M Systems</td><td>719000 A-M Systems</td><td>Use this to make a tool for removing embryos from the brood pouch of anaesthetized females.</td></tr></tbody></table>

ablation of a single cell from eight-cell embryos of the amphipod crustacean parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a promising model organism for laboratory studies of development, providing a useful outgroup comparison to the well studied arthropod model organism Drosophila melanogaster. In contrast to the syncytial cleavages of Drosophila, the early cleavages of Parhyale are holoblastic. Fate mapping using tracer dyes injected into early blastomeres have shown that all three germ layers and the germ line are established by the eight-cell stage. At this stage, three blastomeres are fated to give rise to the ectoderm, three are fated to give rise to the mesoderm, and the remaining two blastomeres are the precursors of the endoderm and germ line respectively. However, blastomere ablation experiments have shown that Parhyale embryos also possess significant regulatory capabilities, such that the fates of blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres. Blastomere ablation has previously been described by one of two methods: injection and subsequent activation of phototoxic dyes or manual ablation. However, photoablation kills blastomeres but does not remove the dead cell body from the embryo. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications. Here we present a protocol for manual removal of single blastomeres from the eight-cell stage of Parhyale embryos, illustrating the instruments and manual procedures necessary for complete removal of the cell body while keeping the remaining blastomeres alive and intact. This protocol can be applied to any Parhyale cell at the eight-cell stage, or to blastomeres of other early cleavage stages. In addition, in principle this protocol could be applicable to early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

Following successful ablation of single third cleavage P. hawaiensis micromeres as described in this protocol, the remaining micromeres gradually shift their positions slightly so as to partially occupy the space formerly occupied by the ablated blastomere. For example, when g is removed, the neighboring blastomeres mr and ml shift slightly and come to share the lateral cell borders that were formerly in direct contact with g (compare&#160;<stro...

Watch this Scientific Journal Video about Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis at JoVE.com

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a promising model organism for studies of crustacean embryology and comparative arthropod development and evolution. This protocol describes a method for manual removal of single blastomeres from early cleavage stage embryos of Parhyale.

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a ...

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Research

JoVE Journal

Biology

12.5K Views.  Harvard University.  De amphipod Parhyale hawaiensis is een veelbelovend model organisme voor studies van schaaldieren embryologie en vergelijkende geleedpotigen ontwikkeling en evolutie. Dit protocol beschrijft een methode voor het handmatig verwijderen van enkele blastomeren van vroeg decollete stadium embryo's van Parhyale. 

Video: Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Developmental Biology

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

Genetics

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Neuroscience

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

In multicellular systems, individual cells are surrounded by the various physical and chemical cues coming from neighboring cells and the environment. This tissue complexity confounds the identification of causal link between extrinsic cues and cellular dynamics. A synthetically reconstituted multicellular system overcomes this problem by enabling researchers to test for a specific cue while eliminating others. Here, we present a method to reconstitute cell contact patterns with isolated Caenorhabditis elegans blastomere and adhesive polystyrene beads. The procedures involve eggshell removal, blastomere isolation by disrupting cell-cell adhesion, preparation of adhesive polystyrene beads, and reconstitution of cell-cell or cell-bead contact. Finally, we present the application of this method to investigate the orientation of cellular division axes that contributes to the regulation of spatial cellular patterning and cell fate specification in developing embryos. This robust, reproducible, and versatile in vitro method enables the study of direct relationships between spatial cell contact patterns and cellular responses.

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

Tissue complexities of multicellular systems confound the identification of causal relationship between extracellular cues and individual cellular behaviors. Here, we present a method to study the direct link between contact-dependent cues and division axes using C. elegans embryo blastomeres and adhesive polystyrene beads.

In multicellular systems, individual cells are surrounded by the various physical and chemical cues coming from neighboring cells and the environment. ...

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...

Deep and Spatially Controlled Volume Ablations using a Two-Photon Microscope in the Zebrafish Gastrula

Morphogenesis involves many cell movements to organize cells into tissues and organs. For proper development, all these movements need to be tightly coordinated, and accumulating evidence suggests this is achieved, at least in part, through mechanical interactions. Testing this in the embryo requires direct physical perturbations. Laser ablations are an increasingly used option that allows relieving mechanical constraints or physically isolating two cell populations from each other. However, many ablations are performed with an ultraviolet (UV) laser, which offers limited axial resolution and tissue penetration. A method is described here to ablate deep, significant, and spatially well-defined volumes using a two-photon microscope. Ablations are demonstrated in a transgenic zebrafish line expressing the green fluorescent protein in the axial mesendoderm and used to sever the axial mesendoderm without affecting the overlying ectoderm or the underlying yolk cell. Cell behavior is monitored by live imaging before and after the ablation. The ablation protocol can be used at different developmental stages, on any cell type or tissue, at scales ranging from a few microns to more than a hundred microns.

Embryonic development requires large-scale coordination of cell motion. Two-photon excitation mediated laser ablation allows the spatially controlled 3-dimensional ablation of large groups of deep cells. In addition, this technique can probe the reaction of collectively migrating cells in vivo to perturbations in their mechanical environment.

Morphogenesis involves many cell movements to organize cells into tissues and organs. For proper development, all these movements need to be tightly ...

Preparation of Single-Somite Explants from Zebrafish Embryos

The body axis of vertebrate embryos is periodically subdivided into 3D multicellular units called somites. While genetic oscillations and molecular prepatterns determine the initial length-scale of somites, mechanical processes have been implicated in setting their final size and shape. To better understand the intrinsic material properties of somites, a method is developed to culture single-somite explant from zebrafish embryos. Single somites are isolated by first removing the skin of embryos, followed by yolk removal and sequential excision of neighboring tissues. Using transgenic embryos, the distribution of various sub-cellular structures can be observed by fluorescent time-lapse microscopy. Dynamics of explanted somites can be followed for several hours, thus providing an experimental framework for studying tissue-scale shape changes at single-cell resolution. This approach enables direct mechanical manipulation of somites, allowing for dissection of the material properties of the tissue. Finally, the technique outlined here can be readily extended for explanting other tissues such as the notochord, neural plate, and lateral plate mesoderm.

We present a protocol for isolating single somites from zebrafish embryos, the dynamics of which can be followed in culture for several hours by fluorescence time-lapse microscopy, thus providing a methodology to quantify tissue-scale shape changes at single-cell resolution.

The body axis of vertebrate embryos is periodically subdivided into 3D multicellular units called somites. While genetic oscillations and molecular ...

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal which tissues descend from each cell of the embryo. Although fate maps are very useful for identifying the precursors of an organ and for elucidating the developmental path by which the descendant cells populate that organ in the normal embryo, they do not illustrate the full developmental potential of a precursor cell or identify the mechanisms by which its fate is determined. To test for cell fate commitment, one compares a cell's normal repertoire of descendants in the intact embryo (the fate map) with those expressed after an experimental manipulation. Is the cell's fate fixed (committed) regardless of the surrounding cellular environment, or is it influenced by external factors provided by its neighbors? Using the comprehensive fate maps of the Xenopus embryo, we describe how to identify, isolate and culture single cleavage stage precursors, called blastomeres. This approach allows one to assess whether these early cells are committed to the fate they acquire in their normal environment in the intact embryo, require interactions with their neighboring cells, or can be influenced to express alternate fates if exposed to other types of signals.

The fate of an individual embryonic cell can be influenced by inherited molecules and/or by signals from neighboring cells. Utilizing fate maps of the cleavage stage Xenopus embryo, single blastomeres can be identified for culture in isolation to assess the contributions of inherited molecules versus cell-cell interactions.

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal  which tissues descend from each cell of the embryo.  Although fate maps ...

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

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